Isolation, identification, characterization, and screening of rhizospheric bacteria for herbicidal activity

  • Charles Oluwaseun Adetunji
  • Julius Kola Oloke
  • Gandham Prasad
  • Oluwasesan Micheal Bello
  • Osarenkhoe Omorefosa Osemwegie
  • Mishra Pradeep
  • Ravinder Sing Jolly


The consistent application of agrochemical herbicides has been reported to impact negatively on human health, environment, and food safety, and facilitated the emergence of weed resistances. Rhizosphere bacteria (RB) of different crops were screened for antagonism against Amaranthus hybridus L. (pigweed) and Echinochloa crus-galli (L.) Beauv. (barnyard grass) using necrosis assay technique. A total of eight rhizosphere bacterial isolates (B1–B8) produced different degrees of leaf necrosis on target weeds with isolate B2 manifesting the most significant necrotic activity. The rhizospheric bacterium (B2) with the highest necrotic activity was identified using 16S rRNA sequencing technique and further investigated. Molecular, morphological, and biochemical characterizations confirmed B2 isolate to be Pseudomonas aeruginosa. On isolation with ethyl acetate, separation, defatting, purification, and flash chromatography, seven different fractions (fraction 1–fraction 7) were obtained out of which fraction 4 showed the highest necrotic activity in necrosis assay experiment. Preparative HPLC of fraction 4 resulted in a pure compound that completely inhibited seed germination and seedling development of pigweed and barnyard grass but remained non-antagonistic to other tested soil fungi used in this study. The result obtained from this present study consequently confirmed the antagonistic behavior of rhizosphere-inhabiting P. aeruginosa to the target weeds and qualified the suitability of bacterium as good alternative source of bioherbicide. Potential herbicidal formulation from P. aeruginosa will help reduce crop loss due to weed challenges while offering a partial solution to the use of agrochemicals and food security.


16S rRNA gene DNA sequencing Deleterious rhizosphere bacteria Phylogenetic tree PHPLC and bioherbicide 


Many microbes are potential alternative sources to synthetic herbicides and demonstrate a naturally specialized protection to a range of crops (Flores-Vargas and O’Hara 2006; Kennedy and Stubbs 2007; Harding and Raizada 2015). Synthetic herbicides which may include glyphosate (Hamid et al. 2011), dicamba (Mark et al. 2007), carfentrazone-ethyl (Baghestani et al. 2007), foramsulfuron (Nurse et al. 2007), isoproturon (Muhammad et al. 2012), 2,4-d sodium salt (Makhan et al. 2013), and organic herbicides such as vinegar, clove oil (Brainard et al. 2013), acetic acid (Ivany 2010), corn gluten meal (Johnson et al. 2013), d-limonene, and pelargonic acid (Allen and Randall 2014; Abouziena et al. 2009) are used in global weed control management. Weed management paradigm in recent decades is changing from conventional practices to the use of environment-friendly classical biological and bioherbicidal weed control approaches in forestry, horticulture, and crop production (Javaid and Adrees 2009; Javaid 2010; Javaid et al. 2010, 2011; Javaid and Ali 2011). The shift according to Xiaoya and Mengmeng (2016) was connected to the residual buildup of human health compromising chemicals in the environment combined with the development of weed resistance to synthetic herbicides. While the application of bioherbicides in both horticulture and crop farming is growing, its widespread use in complementing the conventional methods in an integrated weed management system is environmentally, biologically, technically, and commercially limited (Kremer 2005).

Members of fluorescent pseudomonads are commonly free-living, gram-negative microorganisms found in soils, freshwater, and marine environments. They predominate the plant rhizosphere due to the exudation of organic acids, sugars, and amino acids by the plant roots (Lugtenberg and Dekkers 1999). This group of bacteria which includes Pseudomonas, Rhizobacter, Azobacter, Rugamonas, Serpens, and Mesophilobacter has the capacity to grow on simple media compared to media with a large variety of low molecular weight organic compounds. Additionally, they intimately interact with plant roots where they equally invigorate as well as maximize plant nutrient uptake and consequently promote plant overall healthiness, protection, and soil fertility (Kloepper et al. 1980).

The occurrence of weed-associated bacteria that biologically attack weeds has been reported (Kremer et al. 1990). Several studies described the use of agricultural weed-associated fluorescent rhizobacteria as bioherbicides against dicotyledons (Charudattan 1991; Cattelan et al. 1999) and grassy weeds in cereal crop fields (Gurusiddaiah et al. 1994; Xiaoya and Mengmeng 2016). The use of microorganisms as bioherbicides has distinctively lowered the impact of the weed population with valued attraction over conventional methods that have repercussions on the natural ecosystems (Van Driesche et al. 2010). Crump et al. (1999) reported high degree of specificity for target weeds, little or zero effect on non-target crops or humans, and the effective management of herbicide-resistant weed populations as some of the advantages of bioherbicide broadcast applications in weed control. Additionally, bioherbicides may have both pre- and post-emergence effects on weed species (Bolton and Elliott 1989). One group of microorganisms largely overlooked as biocontrol agents of weeds is the deleterious rhizobacteria (DRB) that colonize plant root surfaces and possess the innate ability to inhibit root and shoot development of weeds (Schippers et al. 1987; Kennedy et al. 1991).

Weeds are common pernicious, fast-growing plants in most parts of Africa especially in the tropical and sub-tropical ecosystems of the world. They made up a major fraction of pest challenges in many African farms, plantations, and orchards where they significantly smothered economic crops while simultaneously threatening yields (Takim and Amodu 2013). Echinochloa crus-galli (L.) Beauv. (barnyard grass) and Amaranthus hybridus L. (pigweed) are some of the major weeds threatening economic crop productions. The production of tomatoes (Adigun and Lagoke 2003), maize (Chikoye et al. 2004), sugar cane (Takim et al. 2015), cowpea (Usman et al. 2002; Adigun et al. 2014), bambara groundnut (Asiwe and Kutu 2007), and rice (Ekeleme et al. 2007; Adeosun et al. 2009; FAOSTAT 2013) in many Nigerian farms and orchards may be depleted by 50–86% by unchecked weed invasion (Adigun et al. 2014). Weeds are noted for their seed resilience or long-term survival in soil, rapid population establishment, ability to occupy conveniently human disturbed areas, regenerative characteristics, reproductive capacity, adaptation to spread, development of resistant phenotypes, and abundant seed production (Poston et al. 2000; Heap 2005).

The study aimed to increase the biological sources of bioherbicides by screening rhizospheric bacteria that have potential herbicidal activity and to characterize such using molecular techniques as the basis for bioherbicide formulations.

Materials and methods

Isolation and culture preparation of rhizospheric bacterial

One gram of soil sample each from the rhizosphere of different crops (Table 1) was collected by a sterilized spatula and conveyed to the laboratory in foil paper. Sample sites include farms at the Nigerian Stored Product Research Institute (NSPRI), Ilorin, and Ladoke Akintola University of Technology (LAUTECH), Ogbomosho, respectively. The soil was then mixed with 10 ml distilled water in a sterile glass test tube, shaken, and filtered through a sterile glass funnel stuffed with wool. The filtrate was subjected to serial dilution and cultured using the pour plate method prior to incubation at 37 °C for 48 h on a Mueller Hinton agar. The plates were then observed for bacterial growth and colony count. Colonies were each later screened for herbicidal activity using diluents and necrotic assay technique. Each colony was sub-cultured into fresh medium maintained on slants of Mueller Hinton agar and preserved at 4 °C in the refrigerator according to Fawole and Oso (2004).
Table 1

Details on the location and plant species from where the isolates used in the present investigation were isolated



Plant species




Piper nigrum L. (pepper)

Ogbomosho, Oyo state



Triticum aestivum L.(wheat)

Ilorin, Kwara state



Zea mays L. (maize)

Ogbomosho, Oyo state



Sorghum bicolor (L.) Moench. (Sorghum)

Ilorin, Kwara state



Oryza sativa L. (Oryza)

Ilorin, Kwara state



Psidium guajava L. (guava)

Ogbomosho, Oyo state



Solanum lycopersicum L. (tomatoes)

Ilorin, Kwara state



Carica papaya L. (pawpaw)

Ogbomosho, Oyo state

Bioherbicidal screening of isolates using leaf necrosis assay

The pure colonies of the isolated RB (B1–B8) were each prepared in Mueller Hinton broth and incubated in a rotary shaker (140 rpm) for 12 h at 27 °C. Detached leaves from the target weeds (E. crus-galli and A. hybridus) were surface-sterilized with ethanol, were rinsed until all traces of ethanol are removed, and were each treated with 1 μl of the cell-free filtrate per one leaf surface. This was later transferred to previously labeled petri plates (in triplicates) containing moistened filter paper and cotton ball. The plates were incubated at 25 °C for 7 days and monitored daily for signs of necrotic lesions.

Biochemical and morphological characterization

The biochemical characterization of all the isolates was done according to Cappuccino and Sherman (1992). The tests conducted were gelatin liquefaction, lipid hydrolysis, starch hydrolysis, casein hydrolysis, indole production, hydrogen sulfide test, oxidase test, catalase test, urease test, denitrification, acid and gas production, and arginine hydrolysis. The bacterial isolates were structurally studied using a light microscope and oil immersion objective (×100), gram stain, and fluorescent tests.

Isolation of genomic DNA

The rhizopheric bacterial isolate B2 with the highest degree of herbicidal activity was selected and primed for identification using 16S ribosomal RNA (rRNA) sequencing technique. The AxyPrep Multisource Genomic DNA Miniprep Kit was used to isolate DNA as described by the manufacturer’s manual. The amount of DNA extracted was electrophoresed on 0.8% agarose gel, and the results obtained were compared with a 1-kb ladder.

PCR amplification and sequencing of 16S rRNA gene from B2 isolate

Polymerase chain reaction (PCR) was performed in a Thermal Cycler. The reaction mixture used consisted of 20 ng of genomic DNA, 2.5 U/50 μl of Taq DNA polymerase, and 5 μl of 10× Taq buffer (100 mM Tris-HCl, 500 mM KCl at pH 8.3), and additionally 200 μM dNTP, 10 pmol each of universal primers (forward primer 27F 5′AGAGTTTGATCCTGGCTCAG3′ and reverse primer 1492R5′TACGGTTACCTTGTTACGACTT3′), and 2.0 mM MgCl2. Amplification process denaturation of the extracted genomic DNA was followed by the annealing of primers at 50 °C for 30 s and extension at 72 °C for 1 and 15 min. Five microliters (5 μl) of the amplified product was analyzed by submarine agarose gel (1.2%) electrophoresis with ethidium bromide at 130 V for 30 min followed by visualization under a Gel Doc/UV transilluminator. This was later gel purified using the Qiagen gel extraction kit and thereafter sequenced with 100 ng/μl of 16S rRNA.

Construction and analysis of RB isolate phylogenetic tree

The 16S rRNA sequences were compared and aligned with sequences deposited in the National Center for Bioinformatics (NCBI) GenBank data base using Basic local Alignment Search Tool (BLAST) (Altschul 1997). The sequences were aligned in CLUSTAL X program, and a phylogenetic tree was constructed by the neighbor-joining method program. The 16S rRNA sequences of bacterial isolate B2 were used for constructing the phylogenetic tree.

Isolation and purification of bioherbicidal substance from RB isolate B2 by HPLC method

Thirty-eight grams per liter (38 g/l) of Mueller Hinton broth was prepared in Milli-Q, beef infusion, casein acid hydrolysate, and starch. This was thereafter sterilized before use as the extraction culture at 30 °C. Isolate B2 was then used to inoculate 25 × 2-l flasks each containing 400 ml of the broth and incubated with a rotary shaker at 200 rpm for 72 h. The broth medium was later centrifuged at 120 rpm for 15 min and the supernatant extracted with ethyl acetate to form an organic layer that was later separated using a separating funnel. The resulting filtrate was dried with sodium sulfate and evaporated at 40 °C using a rotary evaporator to obtain the crude product.

Similarly, the crude extract was defatted in a solution of brine (10 ml), methanol (10 ml), and hexane (20 ml) and stirred for 15 min after which the methanol-water layer was extracted with ethyl acetate. The resulting organic layer was dried with sodium sulfate, evaporated at 40 °C on rotary evaporator, and subjected to flash chromatography. The purified and separated fractions were later tested for herbicidal activity while the fraction with the highest degree of herbicidal activity was analyzed by high-performance liquid chromatography (HPLC) using a C-18 column. Elution was done with acetonitrile/water (90/10 v/v) at a flow rate of 1 ml/min, and detection was done at λ254 while the preparative HPLC was at a flow rate of 3 ml/min (Dueñas et al. 2012).

The seeds of the target weeds (E. crus-galli and A. hybridus) were subjected to viability by water, treated with 15% sodium hypochlorite for 20 min, and rinsed with distilled water (Kordali et al. 2007).

Two layers of filter paper were placed at the base of each clean petri dish (9 cm in diameter) and moistened with 10 ml of distilled water. Fifteen viable seeds of E. crus-galli and A. hybridus were placed on the filter paper, separately for pre-emergence test. Then 2.5, 5, 10, and 15 mg/l of the crude and pure extracts respectively were dropped on the filter paper (Whatman No. 1) according to Kordali et al. (2008). The petri dishes were closed with parafilm to prevent escape of volatile compounds, and incubated at 23 ± 2 °C under 12 h of fluorescent light and relative humidity of 80%. The percentage germination and seedling length were evaluated in a completely randomized setup.

Antimycotic analysis

Six soil fungi comprising Aspergillus flavus Link, Fusarium pallidoroseum (Wollenw.) R.F. Castañeda, P. Oliva, Fresneda & N. Rodr., Aspergillus niger Tiegh, Rhizopus stolonifer (Ehrenb.) Stalpers & Schipper, Saccharomyces cerevisiae, and Fusarium oxysporum E.F. Sm. & Swingle were obtained from the culture repository of the Microbiology Department of NSPRI. The test was carried out by growing each fungal species on potato-dextrose-agar (20 ml) amended with 10 and 100 μg/ml of each of the purified fractions respectively. The plates (five per fungal species) were then inoculated with two 7-day-old colony discs and incubated at 25 °C for 15 days. The antifungal effect of the pure fractions isolated from B2 was evaluated by calculating the percentage of linear growth inhibition as 100(y − x)/y, where y = mean colony diameter of toxin-free cultures and x = mean colony diameter of toxin-containing cultures.

Data analysis

Data were analyzed using SPSS software 21. The null hypothesis of equality of mean effect was tested using the two-way ANOVA table at P = 0.05, and means of significant treatments were separated using Duncan’s multiple range tests. Also, the partial eta squared was used to indicate how much of the total variation in the response was accounted for by the factors and their interactions.


Isolation and preliminary screening of rhizopheric bacteria

The result of the preliminary screening of rhizospheric bacteria isolated from different crops showed that isolate B2 from the rhizosphere of wheat plant produced the best necrotic activity on the leaves of target weeds (A. hybridus and E. crus-galli) with values of 2.5 ± 0.5 and 1.7 ± 0.3 mm, respectively, compared to other tested isolates (P = 0.05) (Fig. 1). Poor necrotic activity was observed for both rhizospheric isolates B5 and B7 (0.2 ± 0.1 mm) on A. hybridus while B3, B5, and B7 had a necrotic value of 0.1 ± 0.01 mm on E. crus-galli compared with the necrotic value observed for the control (P = 0.05). B6 and the control shared the same necrotic value against A. hybridus (P = 0.05).
Fig. 1

Preliminary screening of rhizospheric bacterial isolates on Amaranthus hybridus and Echinochloa crus-galli using leaf necrosis assay. Bars were used to indicate same weeds, but weeds with different superscripts are significantly different at P = 0.05

Characterization and identification of herbicidal active isolate B2

The cultural morphology of rhizosphere-inhabiting bacterium (B2) showed that it had a rod shape, light green, irregular, spreading properties and strong level of fluorescence when exposed to UV light on agar. The genomic DNA of the most active DRB strain B2 had 25,000 bases. 16S rDNA gene sequence was submitted to GenBank ( The 16S rRNA sequences were compared and aligned with sequences deposited in the NCBI GenBank database using BLAST for identification of bacteria. The isolated bacterium was identified as Pseudomonas aeruginosa (B2) with an accession number KF976394. The amplified product of the 1500-bp DNA fragment showed (100%) similarity to P. aeruginosa LMG 1242T (Fig. 2).
Fig. 2

Phylogenetic tree construction of the herbicidally active bacterial isolate made in MEGA 5 software using the neighbor-joining method and an outsource group

Isolation and purification of herbicidal principle from B2

The crude extract obtained from B2 (P. aeruginosa) produced seven different fractions during the preparative HPLC analysis. The retention time for fraction 4 was 9.067 min (Figs. 3 and 4). Fraction 4 however showed the best necrotic activity (2.4 mm for pigweed and 1.4 mm for barnyard grass) on the leaves of the two target weeds compared to the other tested fractions (P ≤ 0.05) (Fig. 5).
Fig. 3

HPLC profile of the crude extract from P. aeruginosa (B2) and its bioherbicidal activities

Fig. 4

HPLC profile of the active purified compound from P. aeruginosa B2 showing bioherbicidal activities

Fig. 5

Herbicidal activity of the fraction from active crude of B2 on Amaranthus hybridus and Echinochloa crus-galli using leaf necrosis assay. Bars were used to indicate same weeds, but weeds with different superscripts are significantly different at P = 0.05

Herbicidal assessment of fraction 4 on seed germination and seedling performance of target weeds

The result showed different degrees of inhibitory effect on seed germination and seedling growth of the target weeds compared with control groups. Concentrations of 10 and 15 mg/dish were completely inhibitory seed germination and seedling growth of all the target weeds. The concentrations 2.5 and 5 mg/dish had varied inhibitory effects on the seeds of A. hybridus (5.0 ± 0.87 to 8.0 ± 0.16%) compared to the control (35.20 ± 1.44%) while a better seed germination inhibition range from 9.0 ± 0.20 to 12.00.87% was observed for E. crus-galli compared to the control (58.9 ± 0.67%). The same trend was also observed for the seedling growth of all target weeds compared with the control (P = 0.05) (Table 2).
Table 2

Inhibitory effects of the purified fraction on seed germination and seedling growth of Amaranthus hybridus and Echinochloa crus-galli

Concentration (mg l−1)

Amaranthus hybridus L.

Echinochloa crus-galli (L.) Beauv.

Germination (%)

Root (mm)

Area-part (mm)

Germination (%)

Root (mm)

Area-part (mm)


8.00 ± 0.16de

14.00 ± 0.14c

21.30 ± 0.31e

12.00 ± 0.87c

21.00 ± 0.33b

48.40 ± 0.28b


5.00 ± 0.89e

8.00 ± 0.23d

16.40 ± 1.02f

9.00 ± 0.20cd

18.00 ± 1.00bc

36.20 ± 0.6c


0.00 ± 0.00f

0.00 ± 0.00e

0.00 ± 0.00g

0.00 ± 0.00f

0.00 ± 0.00f

0.00 ± 0.00g


0.00 ± 0.00f

0.00 ± 0.00f

0.00 ± 0.00g

0.00 ± 0.00f

0.00 ± 0.00f

0.00 ± 0.00g


35.20 ± 1.44b

19.30 ± 0.52b

25.30 ± 0.87d

58.90 ± 0.67a

27.20 ± 0.82a

52.50 ± 1.20a

Values are means ± standard error. Means with different lowercase letters within the same column are significantly different (P = 0.05)

Antimycotic assay

Antimycotic assay with the purified fraction from B2 proved equally insignificant (P = 0.05) against the target soil fungi at concentrations up to 100 μg/ml which proved equally (Table 3).
Table 3

Sensitivity of six soil fungi to the purified fraction from B2



10 μg/ml

100 μg/ml


Aspergillus flavus

2.67 ± 0.115b

7.53 ± 0.252a

Fusarium pallidoroseum

2.4 ± 0.265b

18.77 ± 0.635c

Aspergillus niger

3.5 ± 0.1c

17.5 ± 0.0e

Rhizopus stolonifer

6.3 ± 0.173e

12.83 ± 0.058d

Fusarium oxysporum

5.7 ± 0.173d

11.97 ± 0.462f

Saccharomyces cerevisiae

1.6 ± 0.0a

8.87 ± 0.058b

Values are means ± standard error. Means with different lowercase letters within the same column are significantly different (P = 0.05)


Rhizospheric bacteria from different farm crops across various locations in some parts of Nigeria were randomly investigated for their herbicidal activity, and eight of these isolates from a population of other soil microorganisms were observed to be deleterious to target leaves of A. hybridus and E. crus-galli showing different degrees of necrosis. The isolates with observed necrotic activity may qualify as deleterious rhizospheric bacteria (DBR) as remarked by McPhail et al. (2010). Further investigation involving biochemical, molecular (16S rRNA sequencing technique), and morphological characterization proved that the rhizopheric isolate (B2) with the best necrotic activity per time was P. aeruginosa (Sacchi et al. 2002). This rhizospheric bacterial species was equally implicated in related works by Sessitsch et al. (2004) and McPhail et al. (2010). The reason for the various degrees of necrotic activity observed for the rhizospheric bacterial isolates is unclear; it could be assumed to be due to the genetic disposition of the isolates or a combination of other ecological forces such as local moisture and temperature. Pseudomonas species have been demonstrated in literature as a promising DRB with potential for the biological control of weeds (Kennedy et al. 1991; Flores-Vargas and O’Hara 2006; Kremer and Kennedy 1996; Zermane et al. 2007; Caldwell et al. 2012).

Consequent upon the observed result, P. aeruginosa was processed for the isolation of the active compound(s) that may have been responsible for the necrotic activity. The fraction that was observed to show the highest necrotic activity was later tested on the seed germination and seedling growth of pigweeds and barnyard grass. Complete seed germination and seedling growth inhibition were noted at concentrations of 10 and 15 mg/l, respectively, which may be the lethal dose. The hypersensitivity pattern of the P. aeruginosa-derived fraction 4 noticed during the experimentation suggests an antibiosis type of antagonism (Kaewchai et al. 2009). This confirmed that the fraction 4 derivative of P. aeruginosa was inherently chemically configured for herbicidal activity. While further study is required at characterizing the chemical nature of fraction 4, identifying the role of functional groups as well as molecular weights potentiating herbicidal activity, and developing a potent bioherbicidal formulation (Al-Hinai et al. 2010; Yang et al. 2014). Adetunji and Oloke (2013) and Mejri et al. (2013) have formulated a cell suspension of DRB and pasta granules as effective bioherbicides which precludes the use of pure chemical fractions. Moreover, Pseudomonas fluorescence BRG100 isolated from the rhizosphere of green foxtail in the Brooks of Alberta was reportedly used in controlling green foxtail weeds (Caldwell et al. 2012). Additionally, HPLC application in the processing of derivable crude and its purification was also employed by Zhang et al. (2013) for the isolation as well as elucidation of 4-hydroxy-3-methoxycinnamic acid coupled with two indole derivatives from the fungus Pythium aphanidermatum. The extracted fraction was observed to be non-effective against other non-target soil fungi which further validated its suitability as an environmentally safe bioherbicide (Evidente and Motta 2001).


While the variable phenotypes of the bacterium isolated from the rhizosphere of farm crops was properly identified using molecular technique, their test against pigweed and barnyard grass using leaf necrotic assay showed positivity that validates them as a potentially suitable alternative to mitigating the evidential repercussions of chemical herbicide utility. Furthermore, their test specifically for the non-target effect on soil beneficial fungi underscored the potency of the fraction derived from P. aeruginosa as environmentally safe, target-specific, effective, and cheap and having the potential for commercial formulation.



The authors are grateful to the Council of Scientific and Industrial Research (CSIR), New Delhi, India, and The World Academy of Science (TWAS), Italy, for providing the necessary facilities and opportunity to carry out this research. Special thanks to Mr. Rajul Tomar and the whole staff of Microbial Type Culture Collection and Gene Bank (MTCC), Institute of Microbial Technology, Sector 39A, Chandigarh, India for their contribution to the molecular aspect of this work. Also, I like to appreciate Dr. Adejumo Isaac and Miss Onikanni Olayinka for their input in the statistical analysis.


  1. Abouziena HFH, Omar AAM, Sharma SD, Singh M (2009) Efficacy comparison of some new natural product herbicides for weed control at two growth stages. Weed Technol 23(3):431–437CrossRefGoogle Scholar
  2. Adeosun JO, Dauda CK, Gezui MA, Odunze AC, Amapu IY, Kudp T (2009) On-farm weed management in upland rice in three villages of Katsina State of Nigeria. African Crop Science Society. Afr Crop Sci Conf Proc 9:625–629Google Scholar
  3. Adetunji CO, Oloke JK (2013) Efficacy of freshly prepared pesta granular formulations from the multi-combination of wild and mutant strain of Lasiodiplodia pseudotheobromae and Pseudomonas aeruginosa. Albanian J Agric Sci 12(4):555–563Google Scholar
  4. Adigun JA, Lagoke STO (2003) Assessment of critical period of weed interference in transplanted rainfed and irrigated tomatoes in the Nigerian Northern Guinea Savanna. Niger J Plant Prot 21:89–100Google Scholar
  5. Adigun J, Osipitan AO, Lagoke ST, Adeyemi RO, Afolami SO (2014) Growth and yield performance of cowpea (Vigna unguiculata (L.) Walp) as influenced by row-spacing and period of weed interference in South-West Nigeria. J Agric Sci 6(4):188–198Google Scholar
  6. Al-Hinai AH, Al-Sadi AM, Al-Bahry MAS, Al-said FA, Al-Harthi SA, Deadman ML (2010) Isolation and characterization of Pseudomonas aeruginosa with antagonistic activity against Pythium aphanidermatum. J Plant Pathol 92(3):653–660Google Scholar
  7. Allen VB, Randall GP (2014) Management of vegetation by alternative practices in fields and roadsides. Int J Agron 2014:1–12. doi:10.1155/2014/207828 Google Scholar
  8. Altschul SF (1997) Gapped BLAST & PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 25(17):3389–3402CrossRefPubMedPubMedCentralGoogle Scholar
  9. Asiwe J, Kutu RF (2007) Effect of plant spacing on yield weeds insect infestation and leaf bright of Bambara groundnut. Proc Afr Crop Sci Soc 4:1947–1950Google Scholar
  10. Baghestani MA, Zand E, Soufizadeh S, Bagherani N, Deihimfard R (2007) Weed control and wheat (Triticum aestivum L.) yield under application of 2, 4-D plus carfentrazone-ethyl and florasulam plus flumetsulam: evaluation of the efficacy. Crop Prot 26(12):1759–1764CrossRefGoogle Scholar
  11. Bolton H, Elliott LF (1989) Toxin production by a rhizobacterial Pseudomonas sp. that inhibits wheat root growth. Plant Soil 114(2):269–278CrossRefGoogle Scholar
  12. Brainard DC, Curran WS, Bellinder RR (2013) Temperature and relative humidity affect weed response to vinegar and clove oil. Weed Technol 27(1):156–164CrossRefGoogle Scholar
  13. Caldwell CJ, Hynes RK, Boyetchko SM, Korber DR (2012) Colonization and bioherbicidal activity on green foxtail by Pseudomonas fluorescens BRG100 in a pesta formulation. Can J Microbiol 58(1):1–9CrossRefPubMedGoogle Scholar
  14. Cappuccino JG, Sherman N (1992) Microbiology, a laboratory manual. The Benjamin Cummings Publishing Company Inc., California, p 462Google Scholar
  15. Cattelan AJ, Hartel PG, Fuhrmann JJ (1999) Screening for plant growth-promoting rhizobacteria to promote early soybean growth. Soil Sci Soc Am J 63(6):1670–1680CrossRefGoogle Scholar
  16. Charudattan R (1991) The mycoherbicide approach with plant pathogens. In: TeBeest DO (ed) Microbial control of weeds. Chapman & Hall, New York, pp 24–57. doi:10.1007/978-1-4615-9680-6_2
  17. Chikoye D, Schulz S, Ekeleme F (2004) Evaluation of integrated weed management practices for maize in the northern guinea savanna of Nigeria. Crop Prot 23(10):895–900CrossRefGoogle Scholar
  18. Crump NS, Ash GJ, Fagan RJ (1999) The development of an Australian bioherbicide. 12 Australian Weed Conference, pp 235237Google Scholar
  19. Dueñas M, González-Manzano S, Surco-Laos F, González-Paramas A, Santos-Buelga C (2012) Characterization of sulfated quercetin and epicatechin metabolites. J Agric Food Chem 60(14):3592–3598CrossRefPubMedGoogle Scholar
  20. Ekeleme E, Kamara AY, Oikeh SO, Chikoye D, Omoigui LO (2007) Effect of weed competition on upland rice production in north-eastern Nigeria. Afr Crop Sci Conf Proc 8:61–65Google Scholar
  21. Evidente A, Motta A (2001) Phytotoxins from fungi, pathogenic for agrarian forestal and weedy plants. In: Tringali C (ed) Bioactive compounds from natural source. Taylor& Francis, London, pp 473–525Google Scholar
  22. FAOSTAT (2013) Food and agricultural organization of the United States. Date retrieved, 27th March, 2017.
  23. Fawole MO, Oso BA (2004) Biochemical test. Laboratory manual of microbiology. Spectrum Books Limited, Ibadan, pp 14–17Google Scholar
  24. Flores-Vargas RD, O’Hara GW (2006) Isolation and characterization of rhizosphere bacteria with potential for biological control of weeds in vineyards. J Appl Microbiol 100(5):946–954CrossRefPubMedGoogle Scholar
  25. Gurusiddaiah S, Gealy DR, Kennedy AC, Ogg AG (1994) Isolation and characterization of metabolites from Pseudomonas fluorescens-D7 for control of downy brome (Bromus tectorum). Weed Sci 42:492–501Google Scholar
  26. Hamid AA, Aiyelaagbe OO, Balogun GA (2011) Herbicides and its applications. Adv Nat Appl Sci 5(2):201–213Google Scholar
  27. Harding DP, Raizada MN (2015) Controlling weeds with fungi, bacteria and viruses: a review. Front Plant Sci 6:1–14CrossRefGoogle Scholar
  28. Heap I (2005) The international survey of herbicide resistant weeds. Web page:
  29. Ivany JA (2010) Acetic acid for weed control in potato (Solanum tuberosum L.) Can J Plant Sci 90(4):537–542CrossRefGoogle Scholar
  30. Javaid A (2010) Herbicidal potential of allelopathic plants and fungi against Parthenium hysterophorus—a review. Allelopath J 25(2):331–344Google Scholar
  31. Javaid A, Adrees H (2009) Parthenium management by cultural filtrates of phytopathogenic fungi. Nat Prod Res 23(16):1541–1551CrossRefPubMedGoogle Scholar
  32. Javaid A, Ali S (2011) Herbicidal activity of culture filtrates of Trichoderma spp. against two problematic weeds of wheat. Nat Prod Res 25(7):730–740CrossRefPubMedGoogle Scholar
  33. Javaid A, Shafique S, Shafique S (2010) Herbicidal effects of extracts and residue incorporation of Daturametelagainst partheniumweed. Nat Prod Res 24(15):1426–1437Google Scholar
  34. Javaid A, Shafique S, Shafique S (2011) Management of Parthenium hysterophorus (Asteraceae) by Withania somnifera (Solanaceae). Nat Prod Res 25(4):407–416CrossRefPubMedGoogle Scholar
  35. Johnson WC III, Boudreau MA, Davis JW (2013) Combinations of corn gluten meal, clove oil, and sweep cultivation are ineffective for weed control in organic peanut production. Weed Technol 27(2):417–421CrossRefGoogle Scholar
  36. Kaewchai S, Soytong K, Hyde KD (2009) Mycofungicides and fungal biofertilizers. Fungal Divers 38:25–50.
  37. Kennedy AC, Stubbs TL (2007) Management effects on the incidence of jointed goat grass inhibitory rhizobacteria. Biol Control 40(2):213–221Google Scholar
  38. Kennedy AC, Young FL, Elliott LF, Douglas CL (1991) Rhizobacteria suppressive to the weed downy brome. Soil Sci Soc Am J 55(3):722–727Google Scholar
  39. Kloepper JW, Schroth MN, Miller TD (1980)  Effects of Rhizosphere Colonization by Plant Growth-Promoting Rhizobacteria on Potato Plant Development and Yield. Phytopathology 70(11):1078-1082Google Scholar
  40. Kordali S, Cakir A, Sutay S (2007) Inhibitory effects of monoterpenes on seed germination and seedling growth. Zeitschrift fur Naturforschung (C) 62(3):207–214Google Scholar
  41. Kordali S, Cakir A, Ozer H, Cakmakci R, Kesdek M, Mete E (2008) Antifungal, phytotoxic and insecticidal properties of essential oil isolated from Turkish Origanum acutidens and its three components, carvacrol, thymol and p-cymene. Bioresour Technol 99(18):8788–8795CrossRefPubMedGoogle Scholar
  42. Kremer RJ (2005) The role of bioherbicides in weed management. Biopestic Int 1(3,4):127–141Google Scholar
  43. Kremer RJ, Kennedy AC (1996) Rhizobacteria as biocontrol agents of weeds. Weed Technol 10:601–609Google Scholar
  44. Kremer RJ, Begonia MFT, Stanley L, Lanham ET (1990) Characterization of rhizobacteria associated with weed seedlings. Appl Environ Microbiol 56(4):1649–1655PubMedPubMedCentralGoogle Scholar
  45. Lugtenberg BJJ, Dekkers LC (1999) What make Pseudomonas bacteria rhizosphere competent? Environ Microbiol 1(1):9–13CrossRefPubMedGoogle Scholar
  46. Makhan SB, Simerjit K, Tarundeep K, Tarlok S, Megh S, Amit JJ (2013) Control of broadleaf weeds with post-emergence herbicides in four barley (Hordeum spp.) cultivars. Crop Prot 43:216–222.
  47. Mark R, Behrens NM, Sarbani C, Razvan D, Wen Z, Bradley J, LaVallee PL, Herman TE, Clemente DP (2007) Weeks dicamba resistance: enlarging and preserving biotechnology-based weed management strategies. Science 316(5828):1185–1188. doi:10.1126/science.1141596 CrossRefGoogle Scholar
  48. McPhail KL, Armstrong DJ, Azevedo MD, Banowetz GM, Mills DI (2010) 4-Formylaminooxyvinylglycine, an herbicidal germination arrest factor (GAF) from Pseudomonas rhizosphere bacteria. J Nat Prod 73(11):1853–1857CrossRefPubMedPubMedCentralGoogle Scholar
  49. Mejri D, Gamalero E, Souissi T (2013) Formulation development of the deleterious rhizobacterium Pseudomonas trivialisX33d for biocontrol of brome (Bromus diandrus) in durum wheat. J Appl Microbiol 114(1):219–228. doi:10.1111/jam.12036
  50. Muhammad AS, Muhammad M, Muhammad AUL, Abid N (2012) Efficacy of various herbicides against weeds in wheat (Triticum aestivum L.) Afr J Biotechnol 11(4):791–799. doi:10.5897/AJB11.3274 CrossRefGoogle Scholar
  51. Nurse RE, Hamill AS, Swanton CJ, Tardif FJ, Sikkema PH (2007) Weed control and yield response to foramsulfuron in corn. Weed Technol 21(2):453–458CrossRefGoogle Scholar
  52. Poston DH, Wilson HP, Hines TE (2000) Imidazolinone resistance in several Amaranthus hybridus populations. Weed Sci 48(4):508–513CrossRefGoogle Scholar
  53. Sacchi CT, Whitney AM, Mayer LW, Morey R, Steigerwalt A, Boras A, Weyant RS, Popovic T (2002) Sequencing of 16S rRNA gene: a rapid tool for identification of Bacillus anthracis. Emerg Infect Dis 8(10):1117–1123CrossRefPubMedPubMedCentralGoogle Scholar
  54. Schippers B, Bakker AW, Bakker PAHM (1987) Interactions of deleterious and beneficial rhizosphere microorganisms and the effect of cropping practices. Annu Rev Phytopathol 25:339–358CrossRefGoogle Scholar
  55. Sessitsch A, Gyamfi S, Tscherko D, Gerzabek MH, Kandeler E (2004) Activity of microorganisms in the rhizosphere of herbicide treated and untreated transgenic glufosinate-tolerant and wild type oilseeds rape grown in containment. Plant Soil 266(1):105–116. doi:10.1007/s11104-005-7077-4
  56. Takim FO, Amodu AA (2013) Quantitative estimate of weeds of sugarcane (Saccharum officinarum L.) crop in Ilorin, southern Guinea savannah of Nigeria. Ethiop J Environ Stud Manag 6(6):611–619Google Scholar
  57. Takim FO, Ahmadu MS, Omotosho SB (2015) Efficacy of Ametryn herbicides on weeds of sugarcane in southern guinea savanna zone of Nigeria. Niger J Agric, Food Environ 11(3):147–151Google Scholar
  58. Usman A, Elemo KA, Lagoke STO, Adigun JA (2002) Nitrogen and weed management in maize intercropped with upland rice. J Sustain Agric 21(1):5–16. doi:10.1300/J064v21n01_03 CrossRefGoogle Scholar
  59. Van Driesche RG, Carruthers RI, Center T, Hoddle MS, Hough-Goldstein J, Morin L, Smith DL (2010) Classical biological control for the protection of natural ecosystem. Biol Control 154(S1):2–33Google Scholar
  60. Xiaoya C, Mengmeng G (2016) Bioherbicides in organic agriculture. Horticulturae 2(3):1–10Google Scholar
  61. Yang J, Hong-zhe CAO, Wang WV (2014) Isolation, identification, and herbicidal activity of metabolites produced by Pseudomonas aeruginosa CB-4. J Integr Agric 13(8):1719–1726CrossRefGoogle Scholar
  62. Zermane N, Souissi T, Kroschel JR, Sikora R (2007) Biocontrol of broomrape (Orobanchecrenata Forsk. and Orobanchefoetida Poir.) by Pseudomonas fluorescens isolate Bf7-9from the faba bean rhizosphere. Biocontrol Sci Tech 17(5):483–497Google Scholar
  63. Zhang LH, Zhang JL, Liu YC, Cao ZY, Han JM, Yang J, Dong JG (2013) Isolation and structural speculation of herbicide-active compounds from the metabolites of Pythium aphanidermatum. J Integr Agric 12:1026–1032CrossRefGoogle Scholar

Copyright information

© Springer Science+Business Media Dordrecht 2017

Authors and Affiliations

  • Charles Oluwaseun Adetunji
    • 1
  • Julius Kola Oloke
    • 2
  • Gandham Prasad
    • 3
  • Oluwasesan Micheal Bello
    • 4
  • Osarenkhoe Omorefosa Osemwegie
    • 1
  • Mishra Pradeep
    • 5
  • Ravinder Sing Jolly
    • 5
  1. 1.Department of Biological Sciences, Applied Microbiology, Biotechnology and Nanotechnology LaboratoryLandmark UniversityOmu AranNigeria
  2. 2.Department of Pure and Applied BiologyLadoke Akintola University of TechnologyOgbomosoNigeria
  3. 3.Microbial Type Culture Collection and Gene Bank, CSIR-Institute of Microbial TechnologyChandigarhIndia
  4. 4.Department of Applied ChemistryFederal University Dutsin-MaDutsin-MaNigeria
  5. 5.Department of Bioorganic LaboratoryInstitute of Microbial TechnologyChandigarhIndia

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